Background: Aseptic surgery by definition is surgery performed without contamination or exposure to pathogens. According to the AWR's and the Guide for the Care and Use of Laboratory Animals, all survival surgery should be performed using aseptic procedures, including the use of sterile surgical gloves, masks, sterile instruments, and aseptic techniques.

Policy:Aseptic technique must be applied to all survival surgeries in which amphibians are involved. Understanding, it may be impossible to achieve an aseptic site on the animal, a "clean contaminated" site can be achieved by applying the same aseptic standard to amphibians. Wherever possible, the same standards should apply to all cold-blooded species.

Guidelines: These guidelines are provided to help ensure optimal results for aseptic survival surgery using laboratory amphibians. Please refer to the Frog Oocyte Surgery SOP for more details on that procedure.

Space requirements: Separate spaces should be used for the following activities:

animal preparation,

surgical area,

postoperative recovery

A clean space should be dedicated to surgery and maintained as such during the surgical procedure. Surfaces should be wiped with disinfectant before and after surgery. The surgical area also should be covered with a clean drape or benchcoat paper.

Surgical instruments and supplies must be sterile. The following paragraph describes the sterilization of instruments.

Sterilization of Instruments: Instruments must be cleaned prior to packaging for sterilization. Sterilization indicators must be used inside the surgical packs and autoclave tape placed on the outside of the pack to ensure sterilization has taken place. Sterilization can be achieved by several methods:

Steam: 250°F., 15 p.s.i., for 30 minutes.

Gas-Ethylene oxide:Exposure requires 8-10 hours in a special cabinet (gas autoclave). Plastic implantable materials must be aerated for two to seven days, depending on equipment recommendations.

Liquid-Immersion in:

2% glutaraldehyde plus 7.05% phenol (Sporocidin, Cidex, Sonacide) Instruments must be immersed for ten hours in a 1:10 solution. They must be rinsed in sterile water or saline prior to use.

Chlorine dioxide (Clidox)- Instruments must be immersed for six hours in a 1:5:1 solution and require rinsing in sterile water or saline prior to use.

Physical-Hot beads: Insertion in a bed of hot beads for 10-15 seconds works well for re-sterilization of instruments when performing multiple survival surgeries. However, this method only sterilizes the instrument tips. The instruments must be free of organic material prior to contacting the beads. Instruments also must be allowed to cool to avoid burning tissue. Sterilized instruments must be placed on a sterile drape to reduce the chances of contamination. A new sterile instrument pack must be used after every four to five individual animals.

Animal preparation: Animals should be anesthetized away from the surgical area. Anesthesia is induced by placing the animal in MS222-treated water at room temperature. Before beginning any procedure, confirm that the MS-222 is within the expiration date. Contact the on-call VCS veterinarian for the most appropriate formulation for the particular amphibian species (Xenopus formula: 0.2% MS222, 5 mM Hepes, pH 7.5; Axolotls: 0.1% MS222). Lack of response to toe pinch or righting reflex should be used to determine depth of anesthesia. Once an anesthetic depth has been achieved with MS222, continued anesthesia can be achieved by lowering the body temperature by placing on foil or saran wrap over crushed ice. Never place the animal directly on crushed ice. The animal's skin should be kept moist throughout the procedure by wetting with fresh tank water.

Surgical Procedures: Amphibian surgeries are considered clean-contaminated surgeries. The surgical technique requires asepsis, gentle tissue handling, minimal dissection of tissue, appropriate use of instruments and correct use of suture material. The surgeon must scrub hands and arms with a disinfectant soap and don a mask and sterilized gloves and use sterile instruments. Draping of the patient and full gowning of the surgeon may help reduce the contamination of the surgical field, but are deemed of little practical benefit in most instances.

Postoperative Care: Recovery is monitored by watching for purposeful movements. All animals must be returned to the animal facility within 12 hours. IACUC approval is required for lab housing, any period beyond 12 hours, and is based on scientific justification. Check the animal daily for 3 consecutive days post-operatively to monitor for wound dehiscence and other signs of complications (i.e., lethargy, wound dehiscence, inflammation etc). Report any complications to VCS.

Currently, there is an exception for fish/amphibians to the Post-operative Analgesic Policy, because the efficacy and impact of analgesics are not well defined in the latter species. There is on-going review of this policy as scientific information is gathered supporting the efficacy or lack of impact of analgesics. At such time, this exception will be reassessed and brought before the IACUC.

Record keeping: The records may be kept in the animal room in a notebook through the recovery period and returned to your lab for storage afterwards. The records must contain the following information:

Date of surgery

Species, animal identification

Protocol number

Procedure

Anesthetic, including dose

Analgesic, if given, including dose

Antibiotic, if given, including dose and route

Postoperative monitoring (daily for the first 3 days unless there are complications)

Complications, contact VCS immediately

The cage card should also indicate the date of surgery and procedure performed.